Biological organisms must compactly store and yet efficiently read the huge amounts of genetic information contained in their DNA. Many DNA-based enzymes involved in these processes function as highly processive molecular motors capable of translocating over thousands of base pairs without detaching from the DNA template. These motors face mechanical obstacles to their movement, especially in the highly packed DNA of chromatin, and many of these obstacles are known to be important regulators of gene expression. The broad goal of this proposal is to address the question of how these DNA-based motors deal with these obstacles. DNA in chromatin is highly compact as compared to naked DNA. The primary packing unit of chromatin, the nucleosome, consists of roughly two turns of DNA wrapped around a core histone octamer. The molecular mechanism by which RNA polymerase deals with nucleosomes during transcription is not fully understood at the molecular level and remains one of the most fundamental questions in biology. We propose a unique, single-molecule, biophysical approach to address the question of how RNA polymerase deals with nucleosomes. The proposed method combines optical trapping with nanometer-precision position detection techniques, and complements ongoing biochemical and structural studies. This approach, which has proven to be a powerful tool for the study of transcription on naked DNA, will provide direct measurements and visualization of individual molecular events of transcription in chromatin in vitro. Two specific aims are proposed: (1) mechanical stability of DNA associated with nucleosomes, and (2) transcription through nucleosomes. Aim number 1 determines the strength of histone-DNA interactions by stretching a nucleosomal DNA from end-to-end and measuring the tension required to disrupt the nucleosomes. Aim number 2 makes a direct observation of the fate of a nucleosome during an encounter with a transcribing RNA polymerase by monitoring the movement of single molecules of RNA polymerase during transcription and simultaneously detecting possible nucleosome disruption events. Using these methods, we will determine the effects of histone acetylation and chromatin remodeling complexes on transcription through nucleosomal DNA. This proposed basic research will help to elucidate the mechanisms of transcription in eukaryotes, and will further establish the technical foundation for mechanical studies of other nucleic acid-based molecular motors at the single molecule level.